Platelet-Rich Plasma in Dermatologic Practice 3030662292, 9783030662295

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Platelet-Rich Plasma in Dermatologic Practice Neil S. Sadick Editor

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Platelet-Rich Plasma in Dermatologic Practice

Neil S. Sadick Editor

Platelet-Rich Plasma in Dermatologic Practice

Editor Neil S. Sadick Department of Dermatology Weill Medical College of Cornell University New York, NY USA

ISBN 978-3-030-66229-5    ISBN 978-3-030-66230-1 (eBook) https://doi.org/10.1007/978-3-030-66230-1 © Springer Nature Switzerland AG 2021 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This Springer imprint is published by the registered company Springer Nature Switzerland AG The registered company address is: Gewerbestrasse 11, 6330 Cham, Switzerland

Preface

The field of dermatology, both aesthetic and medical, has been transformed the past few decades with the emergence of non-invasive and minimally invasive procedures for the face and body that can treat a plethora of conditions with minimal downtime and side effects. To name a few, energy-based devices can not only treat inflammatory conditions such as acne, rosacea, vascular conditions, and pigmentary disorders but also rejuvenate the skin. Injectable fillers and toxins can have applications in cosmetic improvement of conditions such as scarring but also prevent and treat the signs of aging. Together with the number of modalities available and the variety of indications they can treat, there is growth and expectation in the patient population for the efficacy and number of treatments they receive. Across all demographics, patients seek from their dermatologist the latest, most advanced, safe, and effective treatments for all their medical and aesthetic concerns. To this end, regenerative approaches procedures such as platelet-rich plasma (PRP) represent the latest cutting-­edge frontier that, when used alone or in combination with other modalities, can accelerate improvement of a variety of conditions in aesthetic and medical dermatology. Isolated PRP is rich in biologically active factors and is responsible for dermal remodeling, stimulation of stem cells, and vascularization. Growth factors delivered with PRP stimulate healing processes, accelerate cellular proliferation, and rejuvenate soft tissues; it is only natural that it would find robust applications for dermatologic conditions. This book is the first of its kind to describe the use of PRP in dermatology. While PRP has been widely used in the field of musculoskeletal and maxillofacial conditions, only recently has its use been introduced and evaluated in dermatology. From a handful of indications and paucity of evidence a few years ago, today there are hundreds of publications that document the efficacy and safety of PRP for treating hair loss, age-related volume loss, scarring, skin rejuvenation, and other dermatologic disorders. As the field is still in its newborn phase, and protocols have yet to be standardized for many of the treatments, the authors of each chapter, all international authorities in their fields, have put an effort to summarize the evidence and provide their standard of care when it comes to using PRP in their patients. Literature review together with materials and descriptions of techniques allow the readers to have an up-to-date understanding of where the field currently stands. It’s a book primarily directed to dermatologists, both novice and experienced, but also to anyone interested in the applications of PRP in dermatology. v

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We appreciate that there are still many controversies and unanswered questions about the use of PRP.  The depth, frequency, and volume of injections, the ideal period between multiple injections, topical vs injectable delivery, combination protocols with other modalities, and the mechanism by which the most beneficial effect is harnessed are still unclear. In this sense, all contributors recognize that the book is the first step in a long road ahead, where answers will be found, protocols will be solidified, and a standard of care will be in place. Nevertheless, PRP has earned its place as one of the most promising modalities in dermatology, enthusiastically embraced by physicians in the field and, we hope, by the readers of this book too. New York, NY, USA

Neil S. Sadick

Contents

1 Biology of Platelet-Rich Plasma����������������������������������������������������������������   1 Javed Shaik, Ronda Farah, and Maria Hordinsky 2 Platelet-Rich Plasma Preparation Methodologies����������������������������������  13 Amelia K. Hausauer 3 Platelet-Rich Plasma for Skin Rejuvenation ������������������������������������������  27 Gabriela Casabona and Kai Kaye 4 Platelet-Rich Plasma for Wound Healing������������������������������������������������  45 Massimo Del Fabbro, Sourav Panda, Giovanni Damiani, Rosalynn R. Z. Conic, Silvio Taschieri, and Paolo D. M. Pigatto 5 Platelet-Rich Plasma for Hair Loss����������������������������������������������������������  71 Aditya K. Gupta, Jeffrey A. Rapaport, and Sarah G. Versteeg 6 PRP for Scarring and Striae ��������������������������������������������������������������������  83 Michelle Henry 7 Platelet-Rich Plasma for Dermal Augmentation of the Face and Body��������������������������������������������������������������������������������������������  93 Hee J. Kim and Noelani E. González 8 Combination Therapies for PRP�������������������������������������������������������������� 103 Suleima Arruda 9 Controversies in PRP�������������������������������������������������������������������������������� 109 Usama Syed and Sachin M. Shridharani Index�������������������������������������������������������������������������������������������������������������������� 117

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Contributors

Suleima Arruda  Suleima Dermatology, Private Practice, San Paolo, Brazil Gabriela  Casabona  Dermatologist and Mohs Surgeon, Scientific Director at Ocean Clinic Marbella, Marbella, Malaga, Spain Rosalynn  R.  Z.  Conic  Young Dermatologists Italian Network (YDIN), Centro Studi GISED, Bergamo, Italy Department of Dermatology, Case Western Reserve University, Cleveland, OH, USA Giovanni  Damiani  Department of Biomedical, Surgical and Dental Sciences, University of Milan, Milan, Italy Clinical Dermatology, IRCCS Istituto Ortopedico Galeazzi, Milan, Italy Young Dermatologists Italian Network (YDIN), Centro Studi GISED, Bergamo, Italy Department of Dermatology, Case Western Reserve University, Cleveland, OH, USA Massimo Del Fabbro  Department of Biomedical, Surgical and Dental Sciences, University of Milan, Milan, Italy Dental Clinic, IRCCS Istituto Ortopedico Galeazzi, Milan, Italy Ronda  Farah  Department Minneapolis, MN, USA

of

Dermatology,

University

of

Minnesota,

Noelani E. González  Icahn School of Medicine at Mount Sinai, New York, NY, USA Aditya K. Gupta  Division of Dermatology, Department of Medicine, University of Toronto School of Medicine, Toronto, ON, Canada Mediprobe Research Inc., London, ON, Canada Amelia  K.  Hausauer  Director of Dermatology, Aesthetx Plastic Surgery and Dermatology, Campbell, CA, USA Michelle  Henry  Cornell Medical College Department of Dermatology, New York, NY, USA Maria  Hordinsky  University of Minnesota, Department of Dermatology, Minneapolis, MN, USA

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Contributors

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Kai  Kaye  Plastic Surgeon, Director at Ocean Clinic Marbella, Marbella, Malaga, Spain Hee J. Kim  Icahn School of Medicine at Mount Sinai, New York, NY, USA Sourav  Panda  Department of Biomedical, Surgical and Dental Sciences, University of Milan, Milan, Italy Department of Periodontics and Oral Implantology, Institute of Dental Science and SUM Hospital, Siksha O Anusandhan, Bhubaneswar, India Paolo  D.  M.  Pigatto  Department of Biomedical, Surgical and Dental Sciences, University of Milan, Milan, Italy Clinical Dermatology, IRCCS Istituto Ortopedico Galeazzi, Milan, Italy Jeffrey A. Rapaport  Cosmetic Skin and Surgery Center, Englewood Cliffs, NJ, USA Javed  Shaik  University Minneapolis, MN, USA

of

Minnesota,

Department

of

Dermatology,

Sachin M. Shridharani  Luxurgery, New York, NY, USA Usama  Syed  Department of Dermatology, Mount Sinai Hospital, New York, NY, USA Silvio  Taschieri  Department of Biomedical, Surgical and Dental Sciences, University of Milan, Milan, Italy Dental Clinic, IRCCS Istituto Ortopedico Galeazzi, Milan, Italy Faculty of Dental Surgery, I. M. Sechenov First Moscow State Medical University, Moscow, Russia Sarah G. Versteeg  Mediprobe Research Inc., London, ON, Canada

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Biology of Platelet-Rich Plasma Javed Shaik, Ronda Farah, and Maria Hordinsky

Basics of Platelet Physiology Platelets comprise 6% of total blood cells, are approximately 2 microns in diameter, and typically number 200,000 to 400,000 per cubic millimeter. Platelets are anucleate derivatives of megakaryocytes in the bone marrow and are formed by the coalescence of cytoplasmic membranes and invaginations of the megakaryocyte surface [1]. It is reported that as many as 100 billion platelets need to be produced daily to maintain an average platelet count of 2–3 × 108 per blood ml [2]. Platelets generally circulate in the blood for 10 days, but when in sites of endothelial injury or disruption, platelets can adhere to a variety of substances including collagen, elastin-­ associated microfibrils, and basement membrane. Platelets also participate in blood-clotting by providing a lipid or lipoprotein surface which induces the conversion of prothrombin to thrombin. More recently, platelets have been implicated in the physiology and pathology of autoimmune disorders [1–3]. Platelets have a multitude of receptors on their cell surface, some of which are upregulated during activation. Platelets in a quiescent state have receptors that help to monitor vascular integrity and upon activation upregulate receptors such as toll-­ like receptors which can recognize pathogens, immune complexes, and siglec receptors which play a role in platelet apoptosis and down-regulation of inflammatory responses [3]. Activated platelets can also release granules of which almost 4000 unique proteins have been identified, with more than 300 of which have been found in platelet lysates. Several of these are megakaryocyte-preformed and are stored in one of three types of granules: dense granules, alpha-granules, or

J. Shaik · M. Hordinsky (*) University of Minnesota, Department of Dermatology, Minneapolis, MN, USA e-mail: [email protected] R. Farah Department of Dermatology, University of Minnesota, Minneapolis, MN, USA © Springer Nature Switzerland AG 2021 N. S. Sadick (ed.), Platelet-Rich Plasma in Dermatologic Practice, https://doi.org/10.1007/978-3-030-66230-1_1

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lysosomes. Dense granules contain ionized calcium, serotonin and adenosine diphosphate, and lysosomes contain a variety of hydrolytic enzymes. Alpha-­ granules contain not only cytokines but also chemokines, growth factors, proteolytic enzymes as well as many other molecules including histamine and antimicrobial peptides [3]. In healthy individuals, approximately 3% of circulating lymphocytes are bound to platelets, and this number significantly increases upon platelet activation. Activated larger lymphocytes have been noted to be more prone to bind to platelets with the following surface proteins contributing to this interaction: P-selectin, GPIIb/IIIa, CD11b, and CD40. Activated platelets can also adhere to circulating neutrophils. Based on these findings, current research in platelet physiology is now focused not only on the role of platelets in blood coagulation but also on their role in autoimmunity.

Platelet-Rich Plasma (PRP) PRP: Background Platelet-rich plasma (PRP) is defined as plasma prepared from autologous blood that contains platelets whose concentration is three- to fivefold above baseline in the blood [4]. In the late 1990s, the beneficial effects of platelet-derived growth factors in healing and tissue regeneration became apparent, and treatment using autologous preparation of PRP was popularized by its use in sports medicine and maxillofacial surgery [5]. PRP is considered to be a safe treatment overall as it is prepared by centrifugation of a patient’s own blood, thereby eliminating any significant risk of an immune reaction. However, there are known clinical risks with the use of PRP in dermatology including but not limited to bleeding, infection, bruising, and pain. Skin necrosis and blindness have also been reported [6]. The PRP preparation process is rapid, requiring minimal specialized equipment and training, thus making PRP an extremely attractive outpatient procedure in the clinical setting. In dermatology, the use of PRP has recently grown exponentially as a popular choice to treat acute and chronic wounds, scars, alopecia [7, 8] as well as in cosmetic dermatology for skin rejuvenation [9, 10]. PRP is considered to be advantageous when compared to other tissue regeneration therapies such as tissue engineering, gene therapy, or cell therapy due to its overall safety, ease of administration, and affordability. PRP therapy is considered to replicate a normal healing environment due to autologous sources of growth factors involved in the healing process. These benefits are considered to outweigh the risks associated with PRP therapy and have attracted its widespread acceptance. Since the role of platelet-derived growth factors in healing has been independently researched previously, clinical studies directly examining clinical outcomes of PRP treatment have preceded studies aimed at understanding the basic science behind mechanism of PRP action. This lack of understanding about the effects of platelets and the combined effect of secreted growth factors following injection may explain the inconsistencies in reporting clinical outcomes within and between studies [11, 12]. Factors leading to variations in quality and quantity of platelets and their

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secretion of growth factors resulting from different methodologies in PRP preparation [13, 14] as well as patient intrinsic factors [15] may also be contributing to inconsistencies in clinical responses reported in the literature. We will outline a general mechanism of action and discuss several key factors to take into consideration for a consistent PRP response.

Preparation and Composition of PRP Several devices are currently approved by the United States Food and Drug Administration (FDA) to prepare PRP. The requirement for initial amounts of blood varies between devices (8–180 ml) and is most likely related to the varying platelet capture efficiencies. These devices typically utilize single or double rounds of centrifugation of anticoagulated blood with or without a density-based separation gel to yield approximately 6–22 ml of PRP. Platelet-poor plasma (PPP) is the top-most layer of plasma following centrifugation which is largely devoid of platelets and serves as a by-product of the PRP preparation process. The bottom layer of plasma after removal of PPP is the PRP that should be enriched in platelets with concentrations ideally three- to fivefold above baseline in the blood. Both PRP and PPP contain the full complement of plasma proteins including those responsible for coagulation. While it is apparent that the platelet concentration in PRP and their secretion of growth factors vary depending on the device [16, 17] and methodology of PRP preparation [15], the choice of anticoagulant used can also affect platelet numbers in the PRP [18]. Leukocytes may be enriched in PRP depending on the type of device used, thereby also affecting levels of growth factors in PRP [16, 19]. Their role in the clinical response is currently unclear with some clinicians strongly opposed to having any leukocytes in PRP preparations used to treat dermatologic conditions. Platelets undergo activation to release growth factors from alpha-granules which sometimes is included in protocols before PRP administration by addition of thrombin and/or calcium chloride. The method chosen to activate PRP has been shown to affect the quantity of growth factors released by platelets [20]. Nonetheless, platelets undergo spontaneous activation following PRP administration from adhesion to matrix proteins and collagen present in the extracellular matrix [21]. In summary, the quality and quantity of platelets in PRP are dependent not only on device and methodology of preparation but also on the anticoagulant used and method chosen to activate PRP.

Growth Factors in PRP Activated platelets secrete hundreds of proteins [22], most of them from alpha-­ granules. These interact with target cells activating intracellular signaling pathways, thereby inducing cellular proliferation, differentiation, migration, matrix remodeling, angiogenesis, chemotaxis, and inflammation [4, 23]. Some of the prominent growth factors, cytokines, and chemokines released from alpha-granules include

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platelet-derived growth factor (PDGF) comprising subunits A and B forming three isoforms, AA, BB, and AB, transforming growth factor beta1 (TGFβ1), platelet factor4 (PF4), epidermal growth factor (EGF), vascular endothelial growth factor (VEGF), fibroblast growth factor-2 (FGF-2), thrombospondin-1 (TSP-1), plasminogen activator inhibitor (PAI-1), hepatocyte growth factor (HGF), and insulin-like growth factor-1 (IGF-1) [4]. Among these, VEGF, PDGF, and FGF-2 can act as angiogenic factors, TGFβ1 promotes matrix remodeling by improving collagen synthesis by stromal cells and regulates cell proliferation, migration, and apoptosis, while EGF and IGF stimulate cellular proliferation, differentiation, and migration [24, 25]. Due to the complex cocktail of growth factors produced by platelets, several factors found in PRP could have antagonistic effects to tissue regeneration and healing. For example, PRP-associated factors such as angiostatin, endostatin, PF4, TSP-1, and PAI-1 can act as negative regulators of angiogenesis [26]. Similarly, metalloproteinase-9 (MMP-9) detected in platelet alpha-granules and cytoplasm [27] could antagonize matrix remodeling and deposition function of TGFβ1 by degrading collagen and other extracellular matrix proteins. Growth factors mentioned above and several others are secreted by platelets within 1 hour of activation and account to 95% of pre-synthesized growth factors contained within the granules [23]. Platelets continue to synthesize and secrete additional growth factors for the remainder of their life span, usually about 7–10 days [23]. Thus, there is four- to sixfold increase in growth factor concentrations of PDGF-BB, TGFβ1, EGF, and VEGF in PRP upon platelet activation compared to whole blood [28]. Growth factor concentration does not appear to be dependent on platelet counts in PRP since a direct correlation between platelet count and growth factor concentration could not be demonstrated [29, 30]. Instead, platelet activation has been noted to have the greatest influence on immediate growth factor release with high concentrations of calcium and thrombin as activators yielding 6- to 8-fold increase in growth factor concentration [31]. In addition, significant variations in growth factor concentrations, particularly FGF-2 and VEGF, have been observed between individuals despite their having very similar platelet counts in PRP [30, 32]. Gender and age differences in growth factor concentrations in PRP have also been reported [33]. Of those tested, EGF, HGF, IGF-1, and PDGF-BB were significantly higher for females, while EGF, IGF-1, PDGF-AB, PDGF-BB, and TGFβ1 achieved significance for people who were 25 years or younger [33]. The increase in growth factor levels in PRP can also vary with the type of device used. While a direct link between growth factor levels and platelet numbers in PRP has not been established, devices that also yield large numbers of leukocytes tend to have higher levels of PDGF-AB, PDFF-BB, and VEGF in PRP [16].

 echanisms of Action and Common Uses of PRP M in Dermatology Use of PRP has recently made swift entries into the worlds of medical and aesthetic dermatology. Its use originated in the field of hematology in the 1970s for the management of thrombocytopenia [5, 34]. Since that time, colleagues in dentistry,

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orthopedics, plastic surgery, urology, gynecology, and ophthalmology have adapted PRP for clinical use [5]. PRP has also been investigated for use in scars including acne scars and burn scars [35–37]. In the field of aesthetic dermatology, PRP has been utilized for rhytides, tissue augmentation, improving skin texture, and skin rejuvenation [38, 39]. In the next section, the possible mechanisms of action for the use of PRP in the management of patients presenting to dermatology clinics with aging skin, scars, alopecia, or wounds will be reviewed.

Skin Rejuvenation Aging skin histologically is characterized by epidermal thinning resulting from slow keratinocyte turnover rate [40]. Atrophy of the dermis is also noticed due to drastically reduced fibroblast numbers in aging skin, leading to reduction in production of collagen and other extracellular matrix proteins [41]. Since the collagen-rich connective tissue produced and maintained by dermal fibroblasts in the human skin provides structural and functional support, these changes lead to thin, structurally weakened skin associated with the appearance of fine wrinkles in naturally aged skin [41]. Maintenance and activation of dermal fibroblasts are essential for rejuvenation of aged skin. Activated PRP has been shown to stimulate proliferation of human dermal fibroblasts and induce their production of type I collagen [42, 43]. Additionally, activated PRP also is associated with increased expression of MMP-1 and MMP-3 proteins in dermal fibroblasts. These MMPs are thought to play a central role in dermal remodeling of aged skin by facilitating removal of fragmented and disorganized collagen fibrils, thereby providing an appropriate foundation for new collagen deposition [41, 44]. Mesotherapy or skin needling technique used to administer PRP has also been shown to increase dermal collagen levels [42]. Autologous PRP application for three sessions at 2-week intervals on the face of healthy volunteers has been reported to result in significant improvement of general appearance, skin firmness-sagging, and wrinkle state [45]. In one study, PRP treatment for six sessions at 2-week intervals resulted in significantly improved skin turgor and increased epidermal and dermal thickness upon evaluation using the Global Aesthetic Improvement Scale and optical coherence tomography [46]. Skin aging is also associated with loss of ability to retain water, thereby resulting in changes in skin turgor, resilience, and pliability. This phenomenon is linked to a marked disappearance of epidermal hyaluronic acid (HA), a glycosaminoglycan, and a predominant component of skin extracellular matrix with a unique capacity to bind and retain water molecules [47]. Other functions of HA include lubrication of joints when present in synovial fluid, regulating several aspects of tissue repair, including activation of inflammatory cells and the response of fibroblasts and epithelial cells to injury. Thus, a potential increase in HA secretion by dermal fibroblasts can improve skin appearance following PRP treatment for aging skin. Among the growth factors secreted by platelets, PRP, PDGF, TGFβ1, VEGF, EGF, HGF, and keratinocyte growth factor (KGF) are known to directly affect keratinocyte and fibroblast proliferation and induce dermal remodeling by stimulating

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synthesis of new collagen, elastin, and glycosaminoglycans [48]. Due to these beneficial effects, several cosmetic products for skin rejuvenation have been touted, including topical formulations with these growth factors [49]. However, PRP treatments are probably more effective than topical formulations as treatments can result in biological synthesis of growth factors within the dermal compartment facilitating their direct interactions with target receptors on cells and potentially yielding a better response in rejuvenating aging skin.

Treating Scars Scars form as a normal biological process during the remodeling stage of wound healing [50]. During this stage, fibroblasts and keratinocytes produce MMPs and tissue inhibitors of MMPs which play a crucial role in extracellular matrix remodeling [51]. An imbalance in their ratio leads to the development of either an atrophic or hypertrophic scar depending on whether the deposition of collagen and other matrix proteins is inadequate or too exuberant, respectively [51]. Atrophic scars are common as a result of severe acne and are characterized by sunken areas in the skin with pitted appearance. PRP when used as an adjunct along with microneedling, fractional laser resurfacing, ablative fractional CO2 laser, or other topical supplemental therapy has been reported to improve the overall clinical response in the management of atrophic acne scars [35, 52, 53]. TGFβ1 released by platelets in PRP may promote fibroblast and myofibroblast differentiation and extracellular matrix deposition, thereby remodeling the extracellular matrix of the atrophic scar. As TGFβ1 enhances scarring due to fibroblast activation and increased collagen deposition, PRP may be contraindicated for use in hypertrophic scars [54].

Wound Healing Platelets are important regulators of homeostasis with a primary function of repairing damaged blood vessels through aggregation causing closure of endothelial and tissue wounds [55]. Platelets are the first cells to arrive at sites of damage in large numbers and play essential roles during the different stages of wound healing: inflammation, cell proliferation, and remodeling. Platelets can interact with keratinocytes during wound healing and regulate their migration by delaying re-­ epithelialization until wound bed preparation is completed [56]. The trapped platelets following PRP treatment of acute or chronic wounds become activated from interaction with extracellular matrix proteins and de-granulate resulting in release of granule contents. The main growth factors currently known to be involved in wound healing are PDGF, TGFβ1, EGF, FGF-2, VEGF, HGF, and IGF-1 [57], which are released by platelets following activation. PDGF is one of the critical growth factors released by platelets that promotes wound healing by inducing

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angiogenesis, formation of fibrous tissue, and re-epithelialization during wound healing [28]. Due to its role in wound healing, recombinant human PDGF-BB has been approved by the FDA to treat diabetic ulcers under its trade name Regranex [58]. Apart for PDGF, VEGF exerts strong paracrine effects on endothelial cells increasing their permeability, growth, and migration, thereby supporting the wound angiogenesis process [58]. EGF and FGF-2, apart from mediating angiogenesis, play essential roles in the cell proliferation stage of wound healing by promoting epithelial, keratinocyte and fibroblast proliferation, differentiation, growth, and migration [50]. TGFβ1 promotes fibroblast and myofibroblast differentiation, extracellular matrix deposition, and scar formation, thereby promoting wound healing during the final remodeling stage [50]. Thus, several of the growth factors released by platelets in PRP play a crucial role in wound healing process.

Alopecia The PRP literature has boasted improvement in the management of alopecia, with most data existing for non-scarring alopecias, primarily androgenetic alopecia [5, 59]. Despite these promising studies, the literature remains conflicted, with a recent systematic review by Lotti et al. calling for additional data focused on clinical applications for androgenetic alopecia to support its use [60]. Limited data suggesting improvement in cicatricial alopecias, namely lichen planopilaris, also exists [61]. Platelets in PRP secrete growth factors that are able to restore cell proliferation and differentiation in mitotically quiescent precursor cells for tissue regeneration. The dermal papilla (DP) is a major component of the hair follicle and plays a key role in morphogenesis and regeneration of hair follicle as well as serving as a reservoir for precursor cells that are essential for hair induction [62]. Activated PRP when applied to human dermal papilla cells (DPCs) obtained from normal human scalp has been found to cause an increase in proliferation of DPCs and enhance their hairinductive activity [63, 64]. Interestingly, higher concentrations of PRP did not increase proliferation of DPCs in those studies suggesting that hair follicle regeneration may be sensitive to concentration of platelets and platelet-derived growth factors in PRP. In addition, activated PRP-mediated anti-apoptotic effects on DPCs through activation of ERK and Akt signaling pathways have been found to prolongate the anagen phase of the hair cycle [63], and several growth factors released by platelets within PRP have been shown to have a positive impact on hair growth. The growth factors in activated PRP are believed to stimulate transition of hair follicles from telogen (resting phase) to anagen primarily through angiogenesis and neovascularization mediated by VEGF, PDGF, EGF, and FGF-2 [65]. Hair follicle induction and prolongation of the anagen phase have also been demonstrated from the synergistic effect of the growth factors PDGF-AA and FGF-2 on DPCs [66], and DPCs themselves produce growth factors such as IGF-1, FGF-7, HGF, and VEGF that are necessary to maintain the hair follicle in the anagen phase of the hair cycle [67].

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Other Potential Uses In other segments of medical and aesthetic dermatology, PRP has also been investigated and reportedly been broadly helpful. Case reports of clinical improvement utilizing PRP for skin ulcerations secondary to diabetes, peripheral artery disease, and polyarteritis nodosa [68, 69] are available. In 2017, a portion of patients treated with PRP for vulvar lichen sclerosus experienced decreased inflammation [70]. The PRP literature is also filled with reports of improvement in clinical outcomes when using PRP as part of a combination treatment protocol. This includes post-fractional resurfacing and microneedling [39]. More recently, combination of microneedling and autologous PRP for management of melasma in 23 patients was reportedly helpful [71]. Microneedling has been combined with PRP in hopes of improved outcomes in those with androgenetic alopecia or alopecia areata [72].

Summary These achievements and reports within the clinical dermatology world are invigorating. The role of key platelet-derived GFs in promoting tissue healing in dermatology as summarized in Table 1.1 is well known. However, the understanding of the clinical applications and basic science studying combined effects of GFs within PRP in dermatology is still in its infancy. The lack of understanding in the lab is complicated by the lack of standardization in the clinical setting. This includes, as Table 1.1  Key platelet-derived growth factors (GFs) in PRP and their major effects on tissue healing in dermatology

Skin rejuvenation

Major effects of PRP Key GFs Dermal remodeling PDGF-AA, -BB, -AB TGFβ1

VEGF FGF-2, KGF (FGF-7) EGF HGF

Mechanism of action Fibroblast activation, proliferation, and migration Chemotactic for immune cells Keratinocyte migration during re-epithelialization Extracellular matrix regeneration Activates fibroblasts leading to type I and type III collagen production Promotes angiogenesis Proliferation and migration of endothelial cells Proliferation of epithelial cells and keratinocytes Proliferation and migration of endothelial cells and fibroblasts Epidermal cell proliferation and migration Extracellular matrix formation and three-dimensional tissue growth

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Table 1.1 (continued)

Wound healing

Major effects of PRP Proliferation

Key GFs PDGF VEGF FGF-2, EGF

Scar management Hair regeneration

Matrix remodeling

TGFβ1

Matrix remodeling

TGFβ1

Dermal papilla cells PDGF-AA, (DPCs) growth FGF-2 VEGF, PDGF, EGF, FGF-2

Mechanism of action Angiogenesis, formation of fibrous tissue, re-epithelialization Endothelial cell permeability, growth, and migration Angiogenesis Proliferation, differentiation, growth, and migration of keratinocytes and fibroblasts Fibroblast and myofibroblast differentiation Extracellular matrix deposition Same as in wound healing DPC growth leading to hair follicle induction and prolongation of anagen Angiogenesis and neovascularization

previously mentioned, variability in clinical devices, platelet activators, platelet numbers, platelet growth factors with growth promoting/inhibiting roles in a given pathological setting, application volume, injection technique, and clinical protocols. While the future of PRP in dermatology is promising, exciting, and overall bright, additional studies are needed to fully understand and optimize its use within the field.

References 1. Patel SR, Hartwig JH, Italiano JE. The biogenesis of platelets from megakaryocyte proplatelets. J Clin Invest. 2005;115:3348–54. 2. Weiss HJ.  Platelet physiology and abnormalities of platelet function. N Engl J Med. 1975;293:580–8. 3. Łukasik Z, Makowski M, Makowska J. From blood coagulation to innate and adaptive immunity: the role of platelets in the physiology and pathology of autoimmune disorders. Rheumatol Int. 2018;38:959–74. 4. Pietrzak WS, Eppley BL. Platelet rich plasma: biology and new technology. J Craniofac Surg. 2005;16:1043–54. 5. Alves R, Grimalt R. A review of platelet-rich plasma: history, biology, mechanism of action, and classification. Ski Appendage Disord. 2018;4:18–24. 6. Kalyam K, Kavoussi SC, Ehrlich M, Teng CC, Chadha N, Khodadadeh S, et al. Irreversible blindness following periocular autologous platelet-rich plasma skin rejuvenation treatment. Ophthalmic Plast Rec. 2017;33:S12–6. 7. Zhang M, Park G, Zhou B, Luo D. Applications and efficacy of platelet-rich plasma in dermatology: a clinical review. J Cosmet Dermatol. 2018;17:660–5. 8. Picard F, Hersant B, Bosc R, Meningaud J. Should we use platelet-rich plasma as an adjunct therapy to treat “acute wounds,” “burns,” and “laser therapies”: a review and a proposal of a quality criteria checklist for further studies. Wound Repair Regen. 2015;23:163–70.

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9. Elghblawi E. Platelet-rich plasma, the ultimate secret for youthful skin elixir and hair growth triggering. J Cosmet Dermatol. 2018;17:423–30. 10. Sclafani AP, Azzi J. Platelet preparations for use in facial rejuvenation and wound healing: a critical review of current literature. Aesthet Plast Surg. 2015;39:495–505. 11. Hussain N, Johal H, Bhandari M.  An evidence-based evaluation on the use of platelet rich plasma in orthopedics – a review of the literature. Sicot-J. 2017;3:57. 12. Giordano S, Romeo M, di Summa P, Salval A, Lankinen P. A meta-analysis on evidence of platelet-rich plasma for androgenetic alopecia. Int J Trichology. 2018;10(1):10. 13. Fitzpatrick J, Bulsara MK, McCrory P, Richardson MD, Zheng M. Analysis of platelet-rich plasma extraction. Orthop J Sports Med. 2017;5:2325967116675272. 14. Amable P, Carias R, Teixeira M, da Pacheco Í, do Amaral R, Granjeiro J, et al. Platelet-rich plasma preparation for regenerative medicine: optimization and quantification of cytokines and growth factors. Stem Cell Res Ther. 2013;4:67. 15. Mazzocca AD, McCarthy MR, Chowaniec DM, Cote MP, Romeo AA, Bradley JP, et  al. Platelet-rich plasma differs according to preparation method and human variability. J Bone Jt Surg. 2012;94:308–16. 16. Castillo TN, Pouliot MA, Kim H, Dragoo JL.  Comparison of growth factor and platelet concentration from commercial platelet-rich plasma separation systems. Am J Sports Med. 2011;39:266–71. 17. Kushida S, Kakudo N, Morimoto N, Hara T, Ogawa T, Mitsui T, et al. Platelet and growth factor concentrations in activated platelet-rich plasma: a comparison of seven commercial separation systems. J Artif Organs. 2014;17:186–92. 18. do Amaral R, da Silva N, Haddad N, Lopes L, Ferreira F, Filho R, et al. Platelet-rich plasma obtained with different anticoagulants and their effect on platelet numbers and mesenchymal stromal cells cehavior in vitro. Stem Cells Int. 2016;2016:7414036. 19. Degen RM, Bernard JA, Oliver KS, Dines JS.  Commercial separation systems designed for preparation of platelet-rich plasma yield differences in cellular composition. HSS J. 2017;13:75–80. 20. Cavallo C, Roffi A, Grigolo B, Mariani E, Pratelli L, Merli G, et  al. Platelet-rich plasma: the choice of activation method affects the release of bioactive molecules. Biomed Res Int. 2016;2016:1–7. 21. Ruggeri ZM, Mendolicchio LG.  Adhesion mechanisms in platelet function. Circ Res. 2007;100:1673–85. 22. Coppinger JA, Cagney G, Toomey S, Kislinger T, Belton O, McRedmond JP, et  al. Characterization of the proteins released from activated platelets leads to localization of novel platelet proteins in human atherosclerotic lesions. Blood. 2004;103:2096–104. 23. Alsousou J, Thompson M, Hulley P, Noble A, Willett K. The biology of platelet-rich plasma and its application in trauma and orthopaedic surgery: a review of the literature. Bone Joint J. 2009;91:987–96. 24. Foster TE, Puskas BL, Mandelbaum BR, Gerhardt MB, Rodeo SA. Platelet-rich plasma. Am J Sports Med. 2009;37:2259–72. 25. Pavlovic V, Ciric M, Jovanovic V, Stojanovic P. Platelet rich plasma: a short overview of certain bioactive components. Open Med-Warsaw. 2016;11:242–7. 26. Peterson JE, Zurakowski D, Italiano JE, Michel LV, Fox L, Klement GL, et al. Normal ranges of angiogenesis regulatory proteins in human platelets. Am J Hematol. 2010;85:487–93. 27. Sheu JR, Fong TH, Liu CM, Shen MY, Chen TL, Chang Y, et al. Expression of matrix metalloproteinase-­9 in human platelets: regulation of platelet activation in in vitro and in vivo studies. Brit J Pharmacol. 2004;143:193–201. 28. Eppley BL, Woodell JE, Higgins J.  Platelet quantification and growth factor analysis from platelet-rich plasma: implications for wound healing. Plast Reconstr Surg. 2004;114:1502–8. 29. Weibrich G, Kleis W, Hafner G, Hitzler WE. Growth factor levels in platelet-rich plasma and correlations with donor age, sex, and platelet count. J Cranio Maxill Surg. 2002;30:97–102.

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30. Martineau I, Lacoste E, Gagnon G.  Effects of calcium and thrombin on growth factor release from platelet concentrates: kinetics and regulation of endothelial cell proliferation. Biomaterials. 2004;25:4489–502. 31. Lacoste E, Martineau I, Gagnon G. Platelet concentrates: effects of calcium and thrombin on endothelial cell proliferation and growth factor release. J Periodontol. 2003;74:1498–507. 32. Cho H, Song I, Park S-Y, Sung M, Ahn M-W, Song K. Individual variation in growth factor concentrations in platelet-rich plasma and its influence on human mesenchymal stem cells. Korean J Lab Med. 2011;31:212–8. 33. Evanson RJ, Guyton KM, Oliver DL, Hire JM, Topolski RL, Zumbrun SD, et al. Gender and age differences in growth factor concentrations from platelet-rich plasma in adults. Mil Med. 2014;179:799–805. 34. Andia I, Abate M.  Platelet-rich plasma: underlying biology and clinical correlates. Regen Med. 2013;8:645–58. 35. Alser OH, Goutos I. The evidence behind the use of platelet-rich plasma (PRP) in scar management: a literature review. Scars Burn Heal. 2018;4:2059513118808773. 36. Ruiz A, Cuestas D, Garcıa P, Quintero J, Forero Y, Galvis I, et al. Early intervention in scar management and cutaneous burns with autologous platelet-rich plasma. J Cosmet Dermatol. 2018;17:1194–9. 37. Aal A, Ibrahim I, Sami N, Kareem I.  Evaluation of autologous platelet-rich plasma plus ablative carbon dioxide fractional laser in the treatment of acne scars. J Cosmet Laser Ther. 2017;20:106–13. 38. Alam M, Hughart R, Champlain A, Geisler A, Paghdal K, Whiting D, et al. Effect of platelet-­ rich plasma injection for rejuvenation of photoaged facial skin: a randomized clinical trial. JAMA Dermatol. 2018;154:1447–52. 39. Sand J, Nabili V, Kochhar A, Rawnsley J, Keller G. Platelet-rich plasma for the aesthetic rurgeon. Facial Plast Surg. 2017;33:437–43. 40. Farage MA, Miller KW, Elsner P, Maibach HI. Characteristics of the aging skin. Adv Wound Care. 2013;2:5–10. 41. Quan T, Fisher GJ. Role of age-associated alterations of the dermal extracellular matrix microenvironment in human skin aging: a mini-review. Gerontology. 2015;61:427–34. 42. Abuaf O, Yildiz H, Baloglu H, Bilgili M, Simsek H, Dogan B. Histologic evidence of new collagen formulation using platelet rich plasma in skin rejuvenation: a prospective controlled clinical study. Ann Dermatol. 2016;28:718–24. 43. Kim D, Je Y, Kim C, Lee Y, Seo Y, Lee J, et  al. Can platelet-rich plasma be used for skin rejuvenation? Evaluation of effects of platelet-rich plasma on human dermal fibroblast. Ann Dermatol. 2011;23:424–31. 44. Quan T, Qin Z, Xia W, Shao Y, Voorhees JJ, Fisher GJ. Matrix-degrading metalloproteinases in photoaging. J Invest Derm Symp Proc. 2009;14:20–4. 45. Yuksel E, Sahin G, Aydin F, Senturk N, Turanli A. Evaluation of effects of platelet-rich plasma on human facial skin. J Cosmet Laser Ther. 2014;16:206–8. 46. Gawdat HI, Tawdy AM, Hegazy RA, Zakaria MM, Allam RS. Autologous platelet-rich plasma versus readymade growth factors in skin rejuvenation: a split face study. J Cosmet Dermatol. 2017;16:258–64. 47. Papakonstantinou E, Roth M, Karakiulakis G. Hyaluronic acid: a key molecule in skin aging. Dermatoendocrinol. 2012;4:253–8. 48. Fabi S, Sundaram H. The potential of topical and injectable growth factors and cytokines for skin rejuvenation. Facial Plast Surg. 2014;30:157–71. 49. Aldag C, Teixeira D, Leventhal PS.  Skin rejuvenation using cosmetic products containing growth factors, cytokines, and matrikines: a review of the literature. Clin Cosmet Investig Dermatol. 2016;9:411–9. 50. Chicharro-Alcántara D, Rubio-Zaragoza M, Damiá-Giménez E, Carrillo-Poveda JM, Cuervo-­ Serrato B, Peláez-Gorrea P, et al. Platelet rich plasma: new insights for cutaneous wound healing management. J Funct Biomaterials. 2018;9:10.

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51. Fabbrocini G, Annunziata M, D’Arco V, Vita DV, Lodi G, Mauriello M, et  al. Acne scars: pathogenesis, classification and treatment. Dermatol Res Pract. 2010;2010:893080. 52. Asif M, Kanodia S, Singh K. Combined autologous platelet-rich plasma with microneedling verses microneedling with distilled water in the treatment of atrophic acne scars: a concurrent split-face study. J Cosmet Dermatol. 2016;15:434–43. 53. Oh I, Kim B, Kim M. Depressed facial scars successfully treated with autologous platelet-rich plasma and light-emitting diode phototherapy at 830 nm. Ann Dermatol. 2014;26:417–8. 54. ris C, Tziotzios C, Vale I. Cutaneous scarring: pathophysiology, molecular mechanisms, and scar reduction therapeutics part I. The molecular basis of scar formation. J Am Acad Dermatol. 2012;66:1–10. 55. Mancuso M, Santagostino E.  Platelets: much more than bricks in a breached wall. Br J Haematol. 2017;178:209–19. 56. Asai J, Hirakawa S, Sakabe J, Kishida T, Wada M, Nakamura N, et al. Platelets regulate the migration of keratinocytes via podoplanin/CLEC-2 signaling during cutaneous wound healing in ice. Am J Pathol. 2016;186:101–8. 57. Grazul-Bilska A, Johnson M, Bilski J, Redmer D, Reynolds L, Abdullah A, et al. Wound healing: the role of growth factors. Drugs Today. 2003;39:787. 58. Murphy PS, Evans GR. Advances in wound healing: a review of current wound healing products. Plast Surg Int. 2012;2012:190436. 59. Hausauer AK, Jones DH. Evaluating the efficacy of different platelet-rich plasma regimens for management of androgenetic alopecia. Dermatol Surg. 2018;44:1191–200. 60. Lotti T, Goren A, Verner I, D’Alessio PA, Franca K. Platelet rich plasma in androgenetic alopecia: a systematic review. Dermatol Ther. 2019:e12837. 61. Bolanča Ž, Goren A, Getaldić-Švarc B, Vučić M, Šitum M. Platelet-rich plasma as a novel treatment for lichen planopillaris. Dermatol Ther. 2016;29:233–5. 62. Driskell RR, Clavel C, Rendl M, Watt FM. Hair follicle dermal papilla cells at a glance. J Cell Sci. 2011;124:1179–82. 63. Li Z, Choi H, Choi D, Sohn K, Im M, Seo Y, et al. Autologous platelet-rich plasma: a potential therapeutic tool for promoting hair growth. Dermatol Surg. 2012;38:1040–6. 64. Xiao S-E, Miao Y, Wang J, Jiang W, Fan Z-X, Liu X-M, et  al. As a carrier–transporter for hair follicle reconstitution, platelet-rich plasma promotes proliferation and induction of mouse dermal papilla cells. Sci Rep. 2017;7:1125. 65. Gupta AK, Carviel J. A mechanistic model of platelet-rich plasma rreatment for androgenetic alopecia. Dermatol Surg. 2016;42:1335–9. 66. Kiso M, Hamazaki TS, Itoh M, Kikuchi S, Nakagawa H, Okochi H.  Synergistic effect of PDGF and FGF2 for cell proliferation and hair inductive activity in murine vibrissal dermal papilla in vitro. J Dermatol Sci. 2015;79:110–8. 67. Jain R, De-Eknamkul W. Potential targets in the discovery of new hair growth promoters for androgenic alopecia. Expert Opin Ther Targets. 2014;18:787–806. 68. Yotsu RR, Hagiwara S, Okochi H, Tamaki T. Case series of patients with chronic foot ulcers treated with autologous platelet-rich plasma. J Dermatol. 2015;42:288–95. 69. Conde-Montero E, Horcajada-Reales C, Clavo P, Delgado-Sillero I, Suárez-Fernández R.  Neuropathic ulcers in leprosy treated with intralesional platelet-rich plasma. Int Wound J. 2016;13:726–8. 70. Goldstein AT, King M, Runels C, Gloth M, Pfau R.  Intradermal injection of autologous platelet-rich plasma for the treatment of vulvar lichen sclerosus. J Am Acad Dermatol. 2017;76:158–60. 71. Hofny ER, Abdel-Motaleb AA, Ghazally A, Ahmed A, Hussein M. Platelet-rich plasma is a useful therapeutic option in melasma. J Dermatol Treat. 2018;29:1–6. 72. Gupta S, Revathi T, Sacchidanand S, Nataraj H.  A study of the efficacy of platelet-rich plasma in the treatment of androgenetic alopecia in males. Indian J Dermatol Venereol Leprol. 2017;83:412.

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Platelet-Rich Plasma Preparation Methodologies Amelia K. Hausauer

Introduction Platelet-rich plasma (PRP) is an autologous solution abstracted from the patient’s own blood containing a small volume of plasma with a concentrated number of platelets and variable other cell lines (leukocytes and erythrocytes). Hence, it reverses the baseline circulating ratio such that platelets constitute roughly 94% and red blood cells less than 5% [1]. However, the precise definition and composition of PRP are controversial and differ by preparation technique. This chapter will discuss the basic steps in PRP preparation, important distinctions in separation techniques, as well as controversies and future directions to better optimize these methodologies.

Definition Originally published in the Oral Maxillofacial and Orthopedic Surgery literature, the classic definition of therapeutic PRP requires a minimum of 1,000,000 platelets/μL, approximately five times that of normal whole blood, based on results from bone and soft-tissue healing studies and in vitro analyses of peak endothelial stimulation and angiogenesis [2, 3]. Others challenge this threshold and propose optimal results that occur in the three- to fivefold range [4, 5], with higher concentrations providing no additional benefit and, in some cases, diminishing the synthetic potential [6–9]. The majority of dermatologic studies fail to report exact parameters of their PRP solution, [10] but among those that do, the mean concentration is roughly threefold [11–29].

A. K. Hausauer (*) Director of Dermatology, Aesthetx Plastic Surgery and Dermatology, Campbell, CA, USA e-mail: [email protected] © Springer Nature Switzerland AG 2021 N. S. Sadick (ed.), Platelet-Rich Plasma in Dermatologic Practice, https://doi.org/10.1007/978-3-030-66230-1_2

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Importantly, obtaining true platelet counts in PRP is not straightforward, so data can be unreliable [30] and cause confusion about appropriate preparation methodologies. Most hematology analyzers are calibrated for whole blood rather than optically lighter platelet products. They may not account for platelet aggregation/ clumping and may have an upper limit for platelet counts above which cells are not recorded [30]. Given these limitations, there is a need for third-party validation and head-to-head concentration studies using appropriate analyzers.

Basic Procedural Steps and Nomenclature Irrespective of system (Table 2.1), all PRP preparation protocols rely on the fact that rapid rotation in a centrifuge layers whole blood into its components based on density, with red cells (RBC) at the base and white blood cells (WBC), platelets, and plasma above. This process involves multiple steps (Figs. 2.1 and 2.2): 1 . Collection of venous whole blood 2. Centrifugation either once (single spin, SS) or twice (double spin, DS) 3. Aspiration of the upper portion of the stratified fluid, which is relatively free of platelets and termed platelet-poor plasma (PPP) 4. Removal of PRP either by • Resuspension of platelets in the residual smaller volume of plasma (mainly in SS systems that contain a gel separator) or • Separation of a buffy coat which contains the majority of platelets and leukocytes sitting just above the RBCs (mainly in DS systems ­ that do not contain a gel separator) 5. Activation of the platelets in PRP endogenously or exogenously (addition of an ‘activator’ in some systems). Table 2.1  PRP preparatory systems used in published clinical studies

Commercial preparatory system (manufacturer) Angel® system (Arthrex Inc) Angel® whole blood separation system (Cytomedix Inc, now Nuo Therapeutics Inc) BTI® system IV (Biotechnology Institute) Cascade-selphyl-esforax system (Aesthetic Factors) CPunT preparation system (Biomed Device) EclipsePRP, also marketed as MyCells (Eclipse MedCorp) GLO PRP centrifuge (Glofinn) GPS III platelet concentration system (Zimmer Biomet) Merisis therapeutics (DiponEd Biointelligence) Omnigrafter (Proteal) P.R.L. platelet rich lipotransfert system (CORIOS Soc. Coop) RegenKit blood cell therapy (RegenLab USA) Smart-PReP 2 system (Harvest Autologous Hemobiologics) Tubex PRP (Moohan Enterprise)

2  Platelet-Rich Plasma Preparation Methodologies Step I

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Step II

Plasma

Step III

PPP Remove top portion of plasma (PPP)

Centrifuge Whole blood and anticoagulant Platelets fibrin

PRP

PRP

Separator gel

Separator gel

Red blood cells white blood cells

Red blood cells white blood cells

Separator gel

Fig. 2.1  Schematic of PRP preparation using a single spin gel separator kit Step I

Step II

Plasma Centrifuge #1

Step III

Collect plasma, buffy coat, discard red blood cells

Step IV

Centrifuge #2

Whole blood and anticoagulant PPP

Red blood cells

Buffy coat (platelets and white blood cells)

Buffy coat + plasma PRP

Fig. 2.2  Schematic of PRP preparation using a double spin buffy coat system

The final solution composition reflects not only individual variations in platelet and growth factor (GF) count among patients but also variations in each preparatory step. For instance, systems differ in the initial volume of whole blood; anticoagulant; time, revolutionary speed (revolutions per minute [rpm] or gravitational force [g]), and number of centrifugations; as well as platelet activation method. Longer or more forceful spin cycles can separate better but may damage platelets, cause premature degranulation, or decrease growth factor viability [2, 31–33]. This variability in technical specifications means that PRP is not a single product but a group of autologous solutions, inconsistently defined in the literature [34, 35]. Some classify PRP into four main categories (Table 2.2): pure PRP (also known as leukocyte-poor PRP, P-PRP); leukocyte-rich PRP (L-PRP); pure platelet-rich fibrin matrix (P-PRF, also known as leukocyte-poor platelet-rich fibrin matrix,

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Table 2.2  PRP nomenclature and categorization [34, 35, 75, 76] Solution characteristics Categorization Pure PRP (P-PRP) Concentrated Leukocyte-poor PRP platelets WBC excluded Suspended in fibrin-rich plasma Leukocyte-rich PRP Concentrated (L-PRP) platelets WBC included Residual RBC Suspended in fibrin-rich plasma Polymerized fibrin Pure platelet-rich clot fibrin matrix Concentrated (PRMF) platelets Platelet rich fibrin WBC variably matrix (PRFM) excluded Polymerized fibrin Leukocyte-, clot platelet-rich fibrin Concentrated (L-PRF) platelets WBC included Forms strong gel membrane

Preparation methods SS: gel separator kits DS: collect PPP with only superficial portion of buffy coat after first spin, centrifuge again, and discard PPP DS: collect PPP with entire buffy coat after first spin, centrifuge again, and discard PPP

DS: spin anticoagulated whole blood, collect PPP plus buffy coat, and activate then incubate until forms a stable clot, special separator gel eliminates WBCs. Note: This forms a stable solid/semi-solid that cannot be injected Solution drawn without any anticoagulant, immediately centrifuged so that central layer forms a clot containing WBC. Note: This forms a stable solid/semi-solid that cannot be injected

Adapted from Eighblawi [75] Leo et al. [76]

PRMF); and leukocyte-rich fibrin and platelet-rich fibrin (L-PRF) [34, 35]. Sadly, published studies do not reliably adhere to this taxonomy, use terms interchangeably, or fail to report parameters in full, making clinical consistency difficult.

System Types There are no standards for the preparation, composition, or administration of platelet-­rich plasma, so evaluating and selecting appropriate systems can be challenging. The first important distinction is between Food and Drug Administration (FDA)-cleared and non-cleared methodologies. Because of its autologous nature, PRP falls under the jurisdiction of the FDA’s Center for Biologics Evaluation and Research (CBER), and preparation systems are cleared as low-risk, 510(k) class II devices for use with bone graft materials in orthopedic procedures. Dermatologic applications are considered off-label. For clearance, systems must demonstrate the ability to concentrate platelets greater than that of whole blood. Other solution characteristics are not monitored explicitly, so there is variability in overall output, variability which may influence clinical outcomes [10, 36]. Centrifuges and in vitro diagnostic (i.e. tiger top or BD®) tubes used for laboratory testing are not FDA

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cleared for PRP production or intended for point-of-care application. Solutions created with these tubes should not be re-injected back into patients, since they are not controlled for endotoxin or pyrogens that may incite potentially serious febrile episodes. Tube labeling states both of these clearly in the “cautions” section of their labeling. Furthermore, these tubes are not intended to separate platelets from other cell lines, and yield is often low, essentially producing PPP [37]. FDA-cleared systems are either manual or automated. Practitioners need to isolate, transfer, and spin the PRP solution at each step of a manual process, while automated systems use closed-loop, computerized methods that are expensive but more sterile and less operator dependent with lower risk of platelet damage and better consistency [2]. Most in-office devices are manual, given the substantial costs (potentially two to ten times more) [38] associated with automated systems, typically found in laboratories, blood banks, or hospitals. Another major distinction is single (SS) versus double (DS) spin based on the number of centrifugation cycles, as noted above. Many DS devices rely on a buffy coat or floating buoy design that produces a higher concentration of not only platelets but also leukocytes and erythrocytes [2, 31–33]. The significance of these other cell lines is discussed in subsequent sections. Small volume, test tube, or gel separator systems include a proprietary anticoagulant gel that helps separate cell lines after only a single spin. Unlike intraoperative applications, which may require large volumes of blood for bone, joint, or wound healing, dermatologists usually provide point-of-care PRP therapy where a compact and reliable small volume system is desirable. Conversely, smaller volumes reduce concentrating capacity with potentially only a few milliliters of yield.

Comparative Analytic Studies: In Vitro and In Vivo Considering this multitude of preparatory methodologies, it is important to define what is being administered and how it may impact clinical outcomes. Most studies come from the orthopedic and oral maxillofacial literature, however, and dermatology-­relevant literature is scarce [9, 35, 39–44]. Kushida et al. characterized PRP composition produced by seven commercially available preparatory systems utilizing a variety of different separation techniques, including SS and DS, tube-­ centrifuged buffy coat, gel separator, and automated [38]. Starting volumes ranged from 8 to 60 mL, and PRP yield was consistently between 0.5 and 3 mL. These volumes are important, since initial blood draw volume impacts feasibility in an outpatient setting and limits concentrating capacity as well as quantity of platelets/ growth factors (GF) delivered to tissue during treatment. In general, DS systems produced solutions with higher platelet, leukocytes, and erythrocyte concentrations, while in SS, gel separator PRP contained significantly fewer WBC and RBC (p